Jiping Pn, Hun Wng, Ye Li*, Chenyng Lu

a School of Marine Science, Ningbo University, Ningbo 315832, China

b Collage of Food and Pharmaceutical Sciences, Ningbo University, Ningbo 315832, China

Keywords:

LC-PUFAs

Fish oil

Krill oil

Label-free quantitative proteomics

Lipids metabolism

A B S T R A C T

Long-chain omega-3 polyunsaturated fatty acids (LC-PUFAs), known for having many health benefits,are usually present in three forms: triglycerides (TG), ethyl esters (EE), and phospholipid (PL). In this study, the effects of these three LC-PUFAs forms (fish oil for TG and EE, krill oil for PL) on the obese mice were compared, and the proteomic changes that focused on lipid metabolism were evaluated via label-free quantitative proteomics analysis. Compared with the model group, all three of the LC-PUFA form supple mentations (labeled as the FO-TG group, FO-EE group and KO-PL groups)could significantly reduce body weight gain (P ＜ 0.01). Low-density lipoprotein cholesterol levels were significantly decreased, whereas high-density lipoprotein cholesterol levels were significantly increased in the FO-TG group and FO-EE group (P ＜ 0.01), and especially in the PL group(P ＜ 0.001). Furthermore, proteomics analysis results suggested that some differentially expressed genes involved in the fatty acid degradation and oxidation pathways had a higher expression fold in the KO-PL group than in the FO-TG or FO-EE groups. Our results showed that dietary LC-PUFAs can reduce fat deposition and inhibit lipogenesis in the liver by upregulating the expression of proteins that are involved in the fatty acid degradation and oxidation pathways. Additionally, KO-PL elicits stronger effects than FO-TG or FO-EE.

1. Introduction

Long-chain omega-3 polyunsaturated fatty acids (LC-PUFAs),such as docosahexaenoic acid (DHA, 22:6n-3) and eicosapentaenoic acid (EPA, 20:5n-3), are well-known for their health benefits. Besides exerting anti-inflammatory properties and preventing the development of heart disease, LC-PUFAs have also been studied as an adjuvant therapy for obesity and its comorbidities [1].

Cold-water marine fish are the main source of omega-3 PUFAs,such as tuna body oils, cod liver oils, amongst others. All these native fish oils exist in the form of triglycerides (TG) with a concentration of approximately 30% DHA and EPA [2]. To increase the concentration of DHA and EPA, TG is usually transferred to ethyl esters (EE), and then molecularly distilled to remove the short chain and saturated fatty acids (SFA). Therefore, in many fish oil products, about 80% –90% and sometimes even higher concentrations of DHA and EPA are bound to EE [3]. Recently, krill oil from Antarctic krill (Euphausia superba), which has a uniquely high content of phospholipid (PL)-bound LC-PUFAs, has garnered attention [4].

The bioavailability of LC-PUFAs in different formulations has been compared in some studies [3,5-7]. Although the LC-PUFAs in krill oil in the form of PL are usually considered to be better absorbed than those in the fish oil in the formulation of EE or TG [6,7],there is still some confusion because of different doses, animals and supplementation time. For example, Lawson and Hughes [8]found that the free fatty acids (FAs) in fish oil were better absorbed (≥ 95% )than the DHA (68% ) and EPA (57% ) in TG, as well as the DHA(20% ) and EPA (21% ) in EE. Dyerberg et al. [2]compared the bioavailability of EE, free FAs and re-esterified TGs against fish oil.Their study demonstrated that compared to natural fish oil, there was 124% bioavailability for re-esterified TGs, 91% for free FAs, and 73% for EE. However, Karin et al. [9]observed similar EPA + DHA levels in the plasms and red blood cells (RBCs) of people whether they consumed fish oil in the form of EE, TG or krill oil in PL, which suggested comparable oral bioavailability irrespective of form.

The effects of LC-PUFAs on the liver are considered to include a decrease in lipogenesis, lesser formation of TGs, and a decreased release of very low-density lipoproteins (VLDLs) into the circulation as well as an increase in FA oxidation [1]. Some studies have indicated that LC-PUFAs inhibit the transcription of genes coding for lipogenesis enzymes, such as FA synthase (FAS) and acetyl-CoA carboxylase (ACC) [10,11]. Meanwhile, LC-PUFAs increased the transcription of regulatory enzymes involved in FA oxidation, such as lipoprotein lipase (LPL) and FA-binding protein (FABP) [11].However, to the best of our knowledge, little is known about the changes mediated by the different forms of LC-PUFAs.

In this study, label-free quantitative proteomics was used to compare the molecular mechanisms by which different forms ofn-3 FAs regulate the expression of enzymes involved in lipid metabolism.Our results contribute to the identification of important mediators involved in the regulation of lipid metabolic pathways.

2. Materials and methods

2.1 Ethics statement

The routine procedures for experimental and animal care in this study were reviewed and approved by the Ningbo University Laboratory Animal Center (affiliated with the Zhejiang Laboratory Animal Common Service Platform), license number SYXK (ZHE 2008 ± 0110).

2.2 Animal and experimental design

Fifty 10-week-old male ICR mice were obtained from the Experimental Animal Breeding Centre, Zhejiang Province, China.The mice were randomly and equally divided into 5 groups(control (C), model (M), FO-TG, FO-EE and KO-PL,n= 10). The mice in the control group were fed a standard chow diet (protein providing 20% kcal, carbohydrate providing 70% kcal, and fat providing 10% kcal) purchased from Laboratory Animal Center of Ningbo University, Ningbo, China. All the mice in the other 4 groups were fed with a high-fat diet (HFD) (66.5% standard chow diet, 10% lard, 1% choline bitartrate, 2.5% cholesterol, and 20% sucrose).Simultaneously, the mice in the experimental groups received natural fish oil in TGs (600 mg/kg·day; FO-TG group), fish oil in EEs(600 mg/kg·day; FO-EE group), and krill oil in PL (600 mg/kg·day;KO-PL group) via oral gavage; the model group received saline via gavage. Throughout the experiment, the mice were kept under standard laboratory conditions as follows: temperature (22 ± 1) °C,dark/light cycles of 12 h/12 h, and relative humidity (55 ± 5)% . The animals had free access to food and water during the study.

The FA compositions of the 3 different formulations (FO-TG,FO-EE and KO-PL) were determined via gas chromatography-mass spectrometry (GC-MS) and are shown in Table 1.

Table 1The FA composition (% ) of 3 different forms of oil in the study.

The mice were kept in the same housing conditions throughout the 12-week duration of the experiments. During the trial, the body weight of each mouse was recorded weekly. At the end of the treatment, mice were anesthetized with 2% sodium pentobarbital(25 mL/kg) and blood samples were collected from the orbital sinus.The blood samples were centrifuged at 3 000 r/min at 4 °C for 15 min to obtain the serum samples.

Serum total cholesterol (TC), TG, low-density lipoprotein cholesterol (LDL-C), and high-density lipoprotein cholesterol(HDL-C) were examined using the respective kits purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China)according to the manufacturer’s instructions.

2.3 Protein preparation and digestion

Mice liver tissues were homogenized in cell lysis buffer (4% SDS,0.1% PMSF and 1 × phosphatase inhibitor), sonicated in an ice bath,and centrifuged at 12 000 ×gfor 40 min to obtain the protein extract.The concentration of the extracted proteins was determined using the BCA assay, followed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The gels were then cut into 1 mm × 1 mm pieces, washed with water, decolorized, reductively alkylated, and enzymatically digested with trypsin (15 h reaction in a 37 °C digester).After the completion of the reaction, proteins were extracted twice by being reacted with 50% acetonitrile and 0.1% tri fluoroacetic acid at 37 °C for 30 min. Finally, the proteins were dried in a freeze-drying apparatus before liquid chromatography-tandem mass spectrometry(LC-MS/MS).

2.4 Peptides identification via LC-MS/MS

Chromatographic separation was performed using ACQUITY UPLC (Waters, USA) with a C18column (3.0 μm, 75 μm × 2.5 cm,Eksigent, USA). The peptides mixture was loaded into the column which was balanced by buffer A (acetonitrile with 0.1% formic acid),and then separated using a linear gradient of buffer B (water with 0.1% formic acid) at a flow rate at 0.3 mL/min.

Spectra scans were acquired using the Q-Exactive mass spectrometry instrument (Thermo Fisher Scientific, USA). MS data was acquired using a data-dependent TOP10 method that dynamically chooses the most abundant precursor ions from the survey scan(350-1 600m/z) for higher-energy collisional dissociation (HCD)fragmentation. Determination of the target value was based on predictive automatic gain control. Survey scans were acquired at a resolution of 70 000 atm/z200 and the resolution for the HCD spectra was set to 17 500 atm/z200.

2.5 Data analysis

The MS data were retrieved using MaxQuant software (Version 1.5.6.5, Germany). The mouse protein library database was used in this study, which contains 16 727 protein sequences and is derived from the UNIPROT database. The data in the MaxQuant search results files were reanalyzed and screened for differential proteins. The search was performed with enzyme specificity trypsin.Meanwhile, a fixed modification of carbamidomethylation of cysteines and a variable modification of oxidation of methionine and protein N-terminal acetylation were included in the database search.The peptides and proteins were identified using a Scaffold assay when they had a probability greater than 95% and 99% probability,respectively.

The protein abundance was calculated with intensity-based absolute quantification (iBAQ) using MaxQuant software. Compared with proteins from the control group, the relative abundance of the identified proteins in the other four groups were calculated and expressed as an “expression fold”. The differentially expressed proteins (DEPs) were those with an expression fold ≥ 2 or ≤ 0.5.Furthermore, gene ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) were employed to determine the biological and functional properties of these DEPs.

2.6 Real-time polymerase chain reaction (PCR)

Total RNA was extracted from mice liver tissues with TRIzol reagent (Takara, Japan). Single-stranded cDNA was synthesized using the reverse transcription kit (Thermo Fisher, USA) according to the manufacturer’s protocol. Brie fly, the genomic DNA was removed using the reaction of incubation at 42 °C for 2 min. Following this, the reverse transcription reaction was initiated and incubation was carried out at 42 °C for 15 min, which was then terminated by heating to 85 °C for 5 min. Actin was used as a reference housekeeping gene and RT-PCR was performed for 4 genes differentially expressed in the different lipid treatments using gene-specific primers (Table 2).RT-PCR was performed using a Rotor-Gene 6 000 RT-PCR PCR detection system (Qiagen, Germany). The conditions of the RT-PCR were as follows: denaturation at 95 °C for 10 min, 40 cycles of 95 °C for 10 s, 60 °C for 15 s, 72 °C for 20 s. Quantification of the PCR product was performed using the ?ΔΔCt method. The data was presented as relative mRNA expression levels (means ± SEM,n= 5) and the results were subjected to pairedt-test analysis. Significant differences between the model group and the other groups were indicated with * forP＜ 0.05, ** forP＜ 0.01 and *** forP＜ 0.001.

Table 2Quantitative RT-PCR primers.

2.7 Statistical analysis

All data were analyzed using the ANOVA and Tukey’s post hoc tests (SPSS, version 19.0, Chicago, IL, USA) and represented as the means ± SEM.P＜ 0.05 was defined as the standard criterion of statistical significance.

3. Results

3.1 Effects of oil supplementation on body weight and blood parameters

The changes in the body weight and blood parameters of the mice that received different formulations of oil are summarized in Table 3.Compared with the model group, PUFA supplementation significantly attenuated body weight gain in all the treatment groups of mice(P＜ 0.01). The serum TC and TG levels of the mice in the FOTG group, FO-EE group and KO-PL groups were all significantly lower than in the model group (P＜ 0.05). The levels of LDL-C were significantly decreased in mice that received KO-PL (P＜ 0.01),and in the FO-TG group (P＜ 0.05), as well as in the FO-EE group(P＜ 0.05) compared to the model group. The opposite trend was observed regarding the levels of HDL-C, which were significantly increased in the mice of the KO-PL group (P＜ 0.001), in the FO-TG and FO-EE group (P＜ 0.01).

Table 3Phenotypic comparison of different forms of oil-fed mice.

3.2 Effects of the different formulations of oil on the proteome of mice liver

Proteins totaling 1 943 were identified in this study via labelingfree proteomic analysis. Proteins featuring a fold change of ≥ 2-fold or ≤ 0.5-fold in each of the two sets of samples were defined as differentially expressed. Compared with the control group, the number of DEPs in the FO-TG, FO-EE, KO-PL and model groups were 225,233, 224 and 204, respectively. As showed in Fig. 1, among the 225 DEPs identified in the FO-TG, 109 proteins were up-regulated and 116 proteins were down-regulated. Moreover, the number of DEPs up-regulated in the FO-EE, KO-PL and the model groups was 124,125 and 109, respectively.

Fig. 1 Numbers of upregulated and downregulated proteins in the liver of mice received LC-PUFAs with different forms.

3.3 Analysis of GO and KEGG

GO annotation was used to determine the molecular function(MF), cell component (CC), and biological process (BP) of all the DEPs (Fig. 2). According to the MF, these proteins were mainly classified into poly(A) RNA binding (GO:00448822), ATP binding(GO:0005524), metal ion binding (GO:0046872), heme binding(GO:0020037), iron ion binding (GO:0005506), structural constituent of ribosome (GO:0003735), arachidonic acid (AA) epoxygenase activity (GO:0008392), calcium binding (GO:0005509), steroid hydroxylase activity (GO:0008395), and fatty-acyl-CoA binding(GO:0000062) proteins. The main CC categories represented were extracellular exosome (GO:0070062), cytoplasm (GO:0005737),mitochondrion (GO:0005739), membrane (GO:0016020), nucleus(GO:0005634), integral component of membrane (GO:16021),endoplasmic reticulum membrane (GO:0005789), endoplasmic reticulum (GO:0005783), extracellular space (GO:0005615), and mitochondrion inner membrane (GO:0005743). While the main BP were oxidation reduction process (GO:0055114), translation(GO:0006412), intracellular protein transport (GO:0006886),positive regulation of transcription (GO:0045944), xenobiotic metabolic process (GO:0006805), exogenous drug catabolic process(GO:0042738), epoxygenase P450 pathway (GO:0019373), response to oxidative stress (GO:0006979), protein transport (GO:0015031),and FA metabolic process (GO:0006631).

Fig. 2 Functional classification and subcellular localization of the DEPs due to dietary with different formulations LC-PUFAs. All the DEPs were categorized according to (A) BP; (B) MF; (C) CC.

Furthermore, these DEPs were also analyzed using the KEGG database and the TOP10 pathways are listed in Table 4. The enriched pathways included steroid hormone biosynthesis (ko00140),retinol metabolism (ko00830), ribosome (ko03010), Alzheimer’s disease (ko05010), FA degradation (ko00071), Parkinson’s disease(ko05012), non-alcoholic fatty liver disease (ko04932), PPAR signaling pathway (ko03320), drug metabolism (ko00983), AA metabolism (ko00590), and chemical carcinogenesis (ko05204).KEGG analysis revealed that most of the metabolic pathways were related to lipid metabolism.

Table 4TOP 10 KEGG pathway enrichment analysis of DEPs.

3.4 The DEPs related to lipid metabolism

The 31 unique DEPs related to lipid metabolism which are also significantly responsive to dietary LC-PUFAs are listed in Table 5.Among those unique DEPs, 9 were associated with FA degradation and oxidation (ABCD3, ACDSB, ACOX1, ACSL1, CPT2, FAAH1,GCDH, PLA2, and THIC), 8 with FA biosynthesis and elongation(ACLY, ECI1, ELOV2, FADS2, FAS, HACD3, HMGCL, and TECR), 3 with AA metabolism (CP3AD, CP4V2, PGES2, and HYEP), 7 with cholesterol metabolism (3BHS7, DHB11, ERLN2,H17B6, NSDHL, SOAT2, and VIGLN), and 4 with the PPAR signaling pathway (ACSF2, ACSM3, APOB, and FABPL).

Table 5DEPs identified in liver tissues of experimental mice compared with model group.

The expression levels of each DEP in every experimental group were analyzed. Relative to the model group, approximately 67% DEPs involved in FA degradation and oxidation were upregulated.In contrast, approximately 78% DEPs involved in FA biosynthesis and elongation were downregulated. Most of the DEPs related to AA metabolism were upregulated, whereas those related to cholesterol homeostasis were downregulated. This suggests that AA metabolism and cholesterol homeostasis are also substantially responsive to dietary LC-PUFAs.

RT-PCR was further used to assess the levels of 6 DEPs genes(Abcd3,Acox1,ApoB,Fads2,Hmgcl, andSoat2) involved in FA metabolism and cholesterol metabolism. As shown in Fig. 3, different mRNA expression patterns were observed for the 6 genes. Except forHmgclandAbcd3,which had opposite regulated trends, the other 4 genes had similar trends with their correspondent proteins.

Fig. 3 RT-PCR analyses of the expression of the unique DEPs in the mice liver which received LC-PUFAs with different formulations. A, Abcd3. B, Acox1.C, ApoB. D, Fads2. E, Hmgcl. F, Soat2.

4. Discussion

In the present work, the effects of three different formulations of LC-PUFAs, i.e. TG, EE and PL, on body weight and blood parameters of HFD-fed obese mice were compared. The results showed that although the total amounts of LC-PUFAs in krill oil ((21.72 ± 0.14)% )are much lower than in fish oils either in EE ((83.51 ± 0.32)% ) or in TG ((42.05 ± 0.42)% ), the effects of krill oil as LC-PUFAs in the PL carrier are more efficient in reducing body weight gain than fish oils(Table 4). This agrees with the results of previous studies, which have reported that the bioavailability of LC-PUFAs in PL formulation is superior, whereas that of fish oil in TG is medium, and that of fish oil in EE is inferior [2]. Another reason for this may be the different amounts of EPA in the different oils. Some reports have shown that the anti-obesity and lipid-lowering effects of EPA are superior to that of DHA [12]. The amount of EPA in KO-PL is (17.82 ± 0.11)% ,which is higher than that in the FO-TG ((5.67 ± 0.05)% ) and FO-EE((8.42 ± 0.11)% ) groups.

To clarify the underlying mechanisms of the different effects of different PUFAs formulations on lipid metabolism, label-free proteomics was used. Compared to the control group, the number of identified DEPs was similar across the three experimental groups (225 for FO-TG, 233 for FO-EE and 224 for KO-PL). In addition, the GO analysis indicated that these proteins were in similar classifications.These results suggested that the metabolic pathways regulated by LCPUFAs are similar regardless of the form.

Furthermore, TOP 10 KEGG pathways enrichment analysis of the DEPs revealed that most of them were related to lipid metabolism,such as steroid hormone biosynthesis, retinol metabolism, FA degradation, etc. Therefore, the expression levels of these DEPs involved in lipid metabolism were further compared among the different experimental groups fed with different forms of LC-PUFAs.

First, most of the proteins in the FA degradation and oxidation pathway were upregulated. It is suggested that the dietary LC-PUFAs inducedβ-oxidation, thus reducing fat deposition and inhibiting of lipogenesis in the liver, which could also mediate the TG-, TC-,LDL-C-lowering and HDL-C-increasing effect of PUFAs [13-15].CarnitineO-palmitoyltransferase 2 (CPT2) and short/branched chain specific acyl-CoA dehydrogenase (ACDSB) are important enzymes of mitochondrial FAβ-oxidation pathways [16]. Compared with the model group, ACDSB and CPT2 in the KO-PL group had the highest expression fold (9.22 for ACDSB and 1.24 for CPT2) than in the FO-TG (6.49 and 1.01, respectively) and FO-EE (2.39 and 0.86,respectively) groups. A similar situation was also observed regarding long chain FA CoA ligase (ACSL) and phospholipase A2 (PLA2),both of which had the highest expression levels in the KO-PL group at 1.60-fold for ACSL and 2.99-fold for PLA2. ACSL and PLA2 are important enzymes involved into the metabolism of PUFAs. PLA2 hydrolyzes membrane phospholipids to release PUFAs and lysophospholipids into the cytosol. The released PUFAs can be re-acylated and incorporated back into cellular phospholipids through the action of ACSL [17]. The increased expression of ACSL and PLA2 suggests that KO-PL has a higher rate of esterification into phospholipids than FO-EE and FO-TG. The higher activity in inducing FAβ-oxidation combined with the higher rate of esterification into phospholipids are considered as the reason why KO-PL appeared more effective than FO-EE and FO-TG in reducing LDL-C and increasing HDL-C levels.

In contrast, most of the DEPs among those involved in the FA biosynthesis and elongation pathways, including elongation of very long chain FAs protein 2 (ELOV2) and FA desaturase 2 (FADS2),were down-regulated. ELOV2 and FADS2 are rate-limiting enzymes,which are responsible for the biosynthesis of FAs, and regulate the specificity of elongation in terms of chain length and degree of unsaturation [18]. In this study, an inhibitory effect of PUFA supplementation in the diet on both ELOV2 and FADS2 was observed and PUFAs with different forms had almost the same responses.

Additionally, the DEPs that involved in the AA metabolism pathway, which were identified in this study and were significantly responsive to dietary PUFAs, were cytochrome P450 3A13 (CP3AD)and prostaglandin E synthase 2 (PGES2). It was observed that both CP3AD and PGES2 were upregulated. AA is released from the phospholipid pools by the activation of phospholipases and free arachidonate is then metabolized along one of three pathways [19].PGES2 is an enzyme involved in AA metabolism, which generates prostaglandin E from prostaglandin H2. CP3AD is an enzyme belonging to the cytochromes P450 family [19]. The upregulation of PGES2 and CP3AD demonstrated that the dietary PUFAs could promote the conversion of AA to other eicosanoids, which have a multitude of potent biological activities. Additionally, the expression folds of both PGES2 and CP3AD were the highest in the FO-EE group than in the other groups. Our results suggested that it is the amount rather than the form of PUFAs that plays a modulator role in AA metabolism.

The repressive expression ofApoBis obvious in obese mice after supplementation with different forms of PUFAs, especially for the KO-PL and FO-TG groups (Table 5 and Fig. 3C). The downregulation ofApoBresults in enhanced lipoprotein lipase (LPL)-mediated catabolism of VLDL and reduced VLDL production, which is considered to be the main mechanism by which PUFAs reduce TG levels [20,21]. It seems that KO-PL and FO-TG, although they contain a lower content of PUFAs, have better effects on reducing TG levels than FO-EE, which contains higher levels of PUFAs.SOAT2 catalyzes the intracellular esterification of cholesterol and selectively inhibits of SOAT2 activity leading to variable decreases in plasma cholesterol content [22]. Our studies indicated that the supplementation with PUFAs downregulates the expression of SOAT2 in obese mice (Table 5). Accordingly, a cholesterol-lowering effect has been observed (Table 3). Although KO-PL led to the lowest TC level (0.67 ± 0.22,P＜ 0.001) in the blood compared to the FO-TG (0.98 ± 0.33,P＜ 0.01) and FO-EE (1.03 ± 0.25,P＜ 0.05)groups, the expression level of SOAT2 in the FO-TG group was the lowest (the expression fold compared with the model group is 0.21),rather than the KO-PL group (the expression fold is 0.84). This result suggests that except for SOAT2, there are other factors affecting cholesterol metabolism when PUFAs are supplied in diet.

Conflict of interest

The authors declare no competing financial interest.

Acknowledgements

This work was supported by the Regional Demonstration Project of Marine Economic Innovation and Development (2013 and 2016),National Natural Science Foundation of China (31800117), and the K.C. Wong Magna Fund offered by the Ningbo University.